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JBIC Journal of Biological Inorganic Chemistry (2021) 26:313–326
https://doi.org/10.1007/s00775-021-01854-y
ORIGINAL PAPER
Conjuring up a ghost: structural and functional characterization
of FhuF, a ferric siderophore reductase from E. coli
I. B. Trindade1 · G. Hernandez1 · E. Lebègue2 · F. Barrière3 · T. Cordeiro1 · M. Piccioli4,5 · R. O. Louro1
Received: 6 December 2020 / Accepted: 23 January 2021 / Published online: 9 February 2021
© The Author(s) 2021
Abstract
Iron is a fundamental element for virtually all forms of life. Despite its abundance, its bioavailability is limited, and thus,
microbes developed siderophores, small molecules, which are synthesized inside the cell and then released outside for iron
scavenging. Once inside the cell, iron removal does not occur spontaneously, instead this process is mediated by siderophoreinteracting proteins (SIP) and/or by ferric-siderophore reductases (FSR). In the past two decades, representatives of the
SIP subfamily have been structurally and biochemically characterized; however, the same was not achieved for the FSR
subfamily. Here, we initiate the structural and functional characterization of FhuF, the first and only FSR ever isolated.
FhuF is a globular monomeric protein mainly composed by α-helices sheltering internal cavities in a fold resembling the
“palm” domain found in siderophore biosynthetic enzymes. Paramagnetic NMR spectroscopy revealed that the core of the
cluster has electronic properties in line with those of previously characterized 2Fe–2S ferredoxins and differences appear
to be confined to the coordination of Fe(III) in the reduced protein. In particular, the two cysteines coordinating this iron
appear to have substantially different bond strengths. In similarity with the proteins from the SIP subfamily, FhuF binds both
the iron-loaded and the apo forms of ferrichrome in the micromolar range and cyclic voltammetry reveals the presence of
redox-Bohr effect, which broadens the range of ferric-siderophore substrates that can be thermodynamically accessible for
reduction. This study suggests that despite the structural differences between FSR and SIP proteins, mechanistic similarities
exist between the two classes of proteins.
Graphic abstract
Extended author information available on the last page of the article
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JBIC Journal of Biological Inorganic Chemistry (2021) 26:313–326
Keywords Ferric-siderophore reductase · Iron uptake · 2Fe–2S protein · Redox-Bohr effect
Abbreviations
ABC ATP-binding cassette
CD Circular dichroism
EPR Electron paramagnetic resonance
FSR Ferric-siderophore reductase
NMR Nuclear magnetic resonance
SAXS Small-angle X-ray scattering
SEC Size exclusion column
SIP Siderophore-interacting protein
TROSY-HSQC Transverse relaxation optimized spectroscopy-heteronuclear single quantum
coherence spectroscopy
Introduction
“Structure without function is a corpse; function without
structure is a ghost”
(Vogel and Wainwright, 1969)
The Great Oxidation Event made the hitherto abundant
iron a trace element as a consequence of precipitation of
iron oxides [1]. Iron has, nonetheless, remained an essential
element for nearly all organisms [2, 3]. To overcome iron
limitation, almost all known bacterial species use siderophores, small molecules that scavenge iron from the extracellular environment, forming Fe(III)-complexes which
are then taken up inside the cell [4, 5]. Different TonBdependent receptors recognize different siderophores into
the periplasm and their transport across the cytoplasmic
membrane is dependent on ABC transporters. Once inside
the cytoplasm, iron can be released via the action of esterases or via reduction by ferric-siderophore reductases [5].
The laboratory strain Escherichia coli K-12 can use diverse
siderophores including hydroxamates (e.g. ferrichrome, ferrioxamine B, coprogen) and catecholates (e.g. enterobactin, yersiniabactin, salmochelin) and it contains at least five
different uptake systems, including the ferrichrome operon
(fhuACDB) and the enterobactin uptake system (fepA, fepB,
fes and fepCDG genes) [6–10]. In the cytoplasm, two distinct ferric-siderophore reductases have been isolated. One
is YqjH that belongs to the SIP (Siderophore-Interacting
Protein) subfamily, which is able to catalyze the release of
iron from Fe(III)-triscatecholates and Fe(III)-dicitrate. The
other is FhuF, from the FSR (Ferric-Siderophore Reductase)
subfamily that showed specificity for a group of hydroxamate-type siderophores, since iron removal from coprogen,
ferrichrome, and ferrioxamine B is significantly reduced
in FhuF-defective mutants [8, 11, 12]. Very little is known
regarding the structure and function of the FSR subfamily
of proteins, given the instability of the pure proteins [11].
13
FhuF is the only protein of the FSR subfamily ever isolated.
Its transcription is derepressed by low iron levels via the iron
regulator Fur, and repressed by OxyR, an oxidative response
regulator [10]. This protein contains a 2Fe–2S cluster with
unusual properties, including the unprecedented binding
motif sequence C–C–x10–C–x2–C, unusual EPR gz value
for this kind of cluster (gz = 1.994), and unusual Mössbauer
parameters with a low quadrupole splitting in the oxidized
form (∆EQ,4.2 K = 0.474 mm s−1) and unusually high quadrupole splitting for the Fe(III) component of the reduced
form (∆EQ190 K = 0.978 mm s−1) [8, 11]. Furthermore, previous studies show that FhuF is truncated at the N-terminal
end, and that it is loosely associated with the cytoplasmic
membrane, since it is possible to purify FhuF from both
cytoplasmic and membrane fractions [11]. Here, we used
a combination of Circular Dichroism (CD), SAXS, Rosetta
modelling, electrochemistry and paramagnetic NMR spectroscopy to advance the structural and functional characterization of FhuF from E. coli K-12.
Materials and methods
Protein production and purification
The plasmid pKF191, derived from pET-19b that codes for
His-tagged FhuF protein was isolated and transformed into
BL21 DE3 competent cells for expression [8, 11]. Freshly
transformed cells were grown in Terrific Broth medium supplemented with 100 mg L−1 ampicillin at 37 ºC, 160 rpm
until they reached an OD of 0.7, the temperature was then
decreased to 30 ºC and cells were collected after 4 h. Cells
were harvested by centrifugation for 10 min at 11,305g and
were then cooled to – 20 ºC. The cells were later defrosted
and resuspended in 20 mM Potassium Phosphate buffer pH
7.6, 300 mM NaCl with a protease-inhibitor cocktail (Roche)
and DNase I (Sigma) prior to a three-pass cell disruption at
6.9 MPa using a French press. The lysate was ultracentrifuged at 204,709g for 90 min at 4 ºC to remove cell membranes and debris. FhuF was purified from the supernatant
using a His-trap affinity column (GE Healthcare) using a
stepwise elution method. The fraction containing FhuF
eluted at 20 mM Potassium Phosphate pH 7.6, 300 mM
NaCl and 250 mM imidazole. Eluted fractions were analyzed by SDS-PAGE with Blue-Safe staining (NZYTech)
and UV–visible spectroscopy to select fractions containing
pure FhuF. The imidazole was removed and FhuF was concentrated at 36 ºC using an Amicon® Ultra Centrifugal Filter
with a cutoff of 10 kDa. For the SAXS data collection, FhuF
fractions were further purified using a Superdex 75 10/300
JBIC Journal of Biological Inorganic Chemistry (2021) 26:313–326
315
GL from GE Healthcare at 1 mL min−1. Samples were kept
at 30 ºC with 0.5 mM of sodium azide and aliquots were
sent for N-terminus sequencing to confirm the identity of
the purified protein.
Circular dichroism
The Far-UV CD spectra of a 6.1 µM FhuF sample in 20 mM
Potassium Phosphate pH 7.6 was recorded on a Jasco-815
spectrophotometer using a 1 mm quartz cell for high performance (QS) (Hellma Analytics). All CD measurements
are an average of four accumulations collected in the
190–260 nm wavelength range using a 0.1 nm data pitch,
and 2 nm bandwidth at 50 nm/min. The CD spectra were
input into the BeStSel webserver to predict secondary elements content [13]. The thermal denaturation of FhuF was
followed by monitoring changes in spectral features as a
function of temperature ranging from 4 to 81 ºC with 7 ºC
steps. The values for the unfolded fraction fU were obtained
by linear extrapolation of the folded 𝜃F and unfolded 𝜃U baselines into the transition zone using the following equation:
fU =
𝜃 − 𝜃F
𝜃U − 𝜃F
(1)
where 𝜃 is the mean residue ellipticity. By fitting to a sigmoidal equation, we extracted the melting temperature (Tm)
of FhuF (i.e., the temperature when fU = 0.5).
Small‑angle X‑ray scattering
Synchrotron SEC-SAXS data on FhuF was collected on the
B21 (ESRF, Grenoble, France) beamline exploiting its inline HPLC system (Agilent 1200 HPLC). To this end, we
injected 50 μL samples with 8.9 mg mL−1 of SEC purified protein in a 4.6 mL Shodex KW402.5-4F size exclusion column at a flow rate of 0.16 mL min−1. Two-second
frames were acquired using a Pilatus 2 M pixel detector.
Data collection conditions are described in Table SI, and
no measurable radiation damage or significant signs of
interparticle interference or aggregation were detected. The
SEC mobile phase consisted of 20 mM potassium phosphate
buffer pH 7.6, 150 mM NaCl. The scattering intensities from
the respective monomeric elution single-peak region were
integrated and buffer subtracted to produce the SAXS-profile
of FhuF using the ScÅtter software [14]. Further processing
was performed using the ATSAS software suite [15]. The P(r)
distribution function was obtained by indirect Fourier Transform. The Rg value was estimated by applying the Guinier
approximation in the range s < 1.3/Rg. The SEC-SAXS
profile raw data were deposited in the repository for smallangle scattering data SASBDB under the project "SAXS of
FhuF—A ferric-siderophore reductase" with the accession
code SASDJ28 [16]. From SEC-SAXS data, a low-resolution
ab initio molecular envelope was generated for FhuF, with
the program DAMMIF using the ATSAS package using the
pair-wise distance distribution (P(r)) calculated from range
0.012 < s < 0.37 Å−1 [15]. Twenty independent models were
generated, and then superimposed and averaged to define
the most populated volume and test the robustness of the
models.
FhuF modeling
The model for FhuF was built by homology modeling using
as templates the known crystal structures of proteins bearing
the FhuF domain, including enzymes involved in iron siderophore biosynthesis in pathogenic bacteria [17]. There are
36 instances of this domain found in the PDB database. The
multi-template modeling was performed with RosettaCM
using evolutionary coupling-derived distance restraints [18,
19]. With RosettaCM, the most homologous portions from
the multiple templates are hybridized into a single model
while modeling the missing residues de novo. We sampled
the diversity of conformational space by building a total of
4000 models, denoted pool. We scored each model by its
Rosetta energy score (Ei) and relative agreement to the SECSAXS data (reduced 𝜒i2) using the following hybrid-scoring
function:
i
i
Zi = ZSAXS
+ ZRosetta
�
�
�
�
𝜒i2 − ⟨𝜒 2 ⟩
�
�
Ei − ⟨E⟩
+ 1 − wsaxs ∙
= wsaxs ∙
𝜎SAXS
𝜎Rosetta
(2)
i
i
where 𝜒i2 and Ei were standardized to ZSAXS
and ZRosetta
,
2
respectively, using the mean values (⟨𝜒 ⟩ and ⟨E⟩) and standard deviations ( 𝜎SAXS and 𝜎Rosetta ) of the pool, with wsaxs
defining the weight of each term (Borges et al., 2020). The
reduced 𝜒i2 was given by CRYSOL 3.0 [15].
NMR spectroscopy
1
H temperature dependence experiments
A sample of approximately 500 μM of oxidized FhuF in
20 mM Potassium Phosphate buffer pH 7.6 with 300 mM
NaCl was lyophilized and solubilized in D2O (99.9 atom %)
for 1H temperature dependence experiments. Reduced FhuF
was obtained using the same sample by degassing it and by
adding an excess of sodium borohydride in an anaerobic
chamber. 1H NMR experiments were performed on a Bruker
Avance II 500 MHz NMR spectrometer equipped with a
5 mm BBI probe. A total of 61,440 transients were acquired
using the super-WEFT pulse sequence (180-τ-90-AQ) with
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JBIC Journal of Biological Inorganic Chemistry (2021) 26:313–326
103 ms of recycle time and τ values of 45 ms, to dampen the
diamagnetic signals and suppress the solvent.
For each temperature, in degrees Celsius, the chemical
shift of each proton signal [(Av/vo)conj] was referenced to
TMS at 0 ppm using the H2O signal as a secondary reference
(5.11–0.012 × T ppm). The temperature dependence of the
contact shift of the cysteine protons was determined according to the Van Vleck formalism:
(
Δv
v0
)con
j
)( �
) ( −Ei )
� ( �
2𝜋g𝜇B Aj ∑ Cji S i S i + 1 . 2S i + 1 exp kT
=
. .
( �
) ( −Ei )
i
3𝛾I kT h
2S + 1 exp
i
kT
(3)
where g is the Free-spin electron g factor, μB is the Bohr
magneton, γI is the free-electron magnetogyric ratio for the
1
H, k is the Boltzmann constant, T is the absolute temperature, Aj is the electron-nuclear coupling constant, h is the
Planck constant, Cji reflect the contribution of each spin (S1
and S2) to the total spin (S’) for each i level [20–23]. The
Ei are the energy values of the eigenstates S’ ranging from
|S1–S2| to |S1 + S2| described by the perturbative Heisenberg
Hamiltonian as a function of the magnetic exchange coupling constant J according to Eq. (4):
Ei =
)
1 �( �
JSi Si + 1
2
(4)
Equation (3) was fit to the experimental data using the
solver routine in MSExcel using standard parameters. Standard errors were determined from the diagonal elements of
the covariance matrix considering 1 ppm experimental
uncertainty in the chemical shift measurements [24].
15 13
N C FhuF: binding experiments
Experiments were performed at 305 K using a Bruker
AVANCE III spectrometer operating at 800 MHz equipped
with a TCI cryoprobe. The reference 1H-15N TROSY-HSQC
experiment was acquired with 64 transients using the pulse
sequence trosyetf3gpsi from the BRUKER catalogue with
250 μM of 15N-13C labeled FhuF in 20 mM Potassium Phosphate buffer pH 7.0 with 300 mM NaCl. The TROSY version of the HSQC was chosen, because it provided sharper
signals that facilitated the analysis. Binding experiments
were performed using samples of 200 μM of 15N-13C labeled
FhuF against increasing amounts of ligand (L): ferrichrome,
apo-ferrichrome. All are oxidized to prevent electron transfer upon binding. Following each addition a 16 transients
1
H-15N TROSY-HSQC experiment was recorded. Chemical
shift perturbations (Δbind) of the NMR signals from FhuF
(Protein, P) were plotted against the molar ratio (R) of [L]/
[P]. The data were fitted using least-squares minimization
to a 1:1 binding model using equations [25]:
13
(
)
√(
)
1 ∞
2
Δ𝛿 bind = Δ𝛿 bind A −
A − 4R
2
A=1+R+
(
)
Kd [P]0 R + [L]0
[P]0 [L]0
(5)
(6)
where Δ𝛿 ∞
is the maximal chemical shift perturbation
bind
of the NMR signals resulting from the complex formation
between the protein and the ligand, and [P]0 is the initial
protein concentration and [L]0 is the stock concentration of
ligand. Only chemical shift perturbations (Δ𝛿 bind ) equal to
or larger than 0.025 ppm were considered significant. The
standard deviation of the fitted value of Kd was calculated
using all data.
Electrochemical experiments
Cyclic voltammetry was performed in a three-electrode cell
with an edge plane pyrolytic graphite disk electrode (PGE,
3 mm diameter) obtained from IJ Cambria Scientific Ltd. as
the working electrode. All cyclic voltammetry measurements
were recorded using an Ag/AgCl, KCl 3 M as reference electrode and a graphite rod as counter electrode. Electrochemical experiments were performed at room temperature with
a SP-300 potentiostat driven by EC-Lab V10.40 software
from Bio-Logic (Bio-Logic Science Instruments, France).
PGE was immersed in diluted protein solution (100 µM) for
3 days before electrochemical measurements in protein-free
aqueous electrolyte (20 mM potassium phosphate buffer) at
different pH values between pH 5 and pH 9. All solutions
were degassed by bubbling argon for 10 min before each
measurement. To facilitate the reading, the potentials discussed in the manuscript are reported in V vs SHE (E/V vs
SHE = E/V vs Ag/AgCl + 0.210 V).
Results
FhuF is a globular helix‑bundle protein
Expression and purification of FhuF gave rise to a folded and
soluble functional domain encompassing residues 18–262.
The protein eluted from the SEC column at a volume corresponding to an apparent molar mass of ca 26.9 kDa,
demonstrating that FhuF (28 kDa) is monomeric, migrating through the column as a folded protein. NMR data in
solution revealed that FhuF is indeed well-folded at 32 ºC
(Fig. 1a) as seen from the broad resonance dispersion of the
1
H–15N fingerprint, reflecting the presence of secondary and
tertiary structure elements, thus compatible with well-folded
protein. The CD profile of FhuF also indicates the presence
JBIC Journal of Biological Inorganic Chemistry (2021) 26:313–326
317
Fig. 1 FhuF is a folded protein. a 2D 1H15N TROSY-HSQC spectrum
of 15N/13C-labeled FhuF collected at 305 K on a Bruker Avance III
800 MHz spectrometer. The resonance map has the typical fingerprint
of a folded protein with large chemical shift dispersion. b CD spectra
of FhuF at different temperatures ranging from 4 and 81 ºC with successive 7 ºC intervals. (Inset) Thermal denaturation as measured by
changes in the signal at 211 and 222 nm
of stable structural elements within the overall protein structure (Fig. 1b), displaying positive values below 200 nm and
two negative bands at 208 and 222 nm, commonly associated with α-helical conformations. BeStSel predicts a helical
content of 55% and 40% of unordered/turn elements with a
small contribution of antiparallel β-sheet [13]. The thermal
denaturation curve of FhuF followed by the changes in the
CD signals at 211 and 222 nm show a cooperative foldingunfolding transition with a Tm of 58–60 ºC, indicating that
the protein has a defined tertiary structure (Fig. 1b, inset)
that unfolds by increasing temperature.
We have employed SAXS to probe further the overall
structure of FhuF. Our synchrotron SEC-SAXS data confirmed that FhuF is also a monomeric globular particle in
solution, with a radius of gyration of 21.5 ± 0.10 Å and a
maximum distance of 88.0 ± 5 Å (Table 1) (Fig. 2a).
The Kratky representation of the protein SAXS data
(Figure S1) is bell-shaped, as expected for highly spherical/globular proteins, contrary to disordered proteins that
do not display a clear maximum [27]. Nevertheless, the
smooth asymmetrical tailing in its pairwise distance distribution (P(r)) suggests the presence of moderate flexibility or a slight deviation from a compact sphere (Fig. 2a).
Accordingly, the SAXS-derived low-resolution structure
is oblong with a spatially separated lobe and two regions
inside the envelope with lower density probability resembling small cavities. To gain more structural insight into
FhuF, we used Rosetta to predict its potential structure,
using multiple templates sharing sequence similarity with
Table 1 SEC-SAXS data analysis
FhuF
Overall parameters
Rg (Å) [from P(r)]
Rg (Å) [from Guinier]
Dmax (Å)
Porod volume estimate, Vp (Å3)
Molecular weight estimate (kDa)a
Oligomeric state
Software
SEC-SAXS data integration
P(r)
Ab initio modelling/ < NSD > (Å)b
Simulated SAXS
SASBDB accession code
21.49 ± 0.10
21.51 ± 0.10
88.0 ± 5.0
37,902.8
25.8 (7.8%)
Monomer
ScÅtter
GNOM 5.0
DAMMIFc/0.727 ± 0.049 Å
CRYSOL
SASDJ28
a
Calculated with SAXSMoW 2.1[26]. The discrepancy to the
sequence weight of FhuF∆1–17 is given inside parentheses
b
c
Mean ± std
Refinement with DAMMIF [15]
FhuF, and incorporating evolutionary coupling constraints
into the homology modeling protocol [18, 19]. Interestingly,
all identified templates were enzymes involved in the iron
siderophore biosynthesis process with a FhuF-like domain
within their overall structure. Examples are the AcsD from
the plant pathogen Pectobacterium chrysanthemi and IucA
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JBIC Journal of Biological Inorganic Chemistry (2021) 26:313–326
Fig. 2 Structural model of FhuF. a SAXS intensity of SEC-purified
FhuF (gray circles), I(s), is represented in logarithmic scale as a function of the momentum of transfer, s. The dark blue line corresponds
to the scattering profile calculated from the ab initio model that best
fitted the experimental data (χ2 = 1.3). Point-by-point deviations of
the fitting are in the bottom panel. The top inset shows the SAXSgenerated ab initio envelope obtained by clustering and averaging 20
independent models, with a Normalized Spatial Discrepancy (NSD)
of 0.727 Å. The bottom inset shows the pair-wise distance distribu-
tion (P(r)) of FhuF. The derived Rg and Dmax values are displayed in
dashed lines. b Rosetta energy vs. SAXS discrepancy, χ2 scatter plot
for the 4000 structure-predictions of FhuF. REU stands for Rosetta
Energy Units. Inset shows expanded scale to highlight the 20 models with the lowest scoring in orange dots, and the best-model as a
purple star. c SAXS ab initio reconstruction of FhuF (blue envelope)
containing the three best-scored homology models. The regions with
high uncertainty are transparent
from the human pathotype Klebsiella pneumonia [17, 28].
Both have three domains that resemble a cupped hand and
these are designated thumb domain 1, palm domain 2, and
fingers domain 3. The palm domain is FhuF-like, and in the
context of AcsD/IucA, contributes with the active site residues. We only used the palm-domain regions as templates.
To improve the modeling and eliminate false-positives, we
used SAXS to score, discriminate, and validate all FhuF
models. SAXS-based approaches were successfully applied
in loop modeling, distinguishing protein–protein interfaces
as well as improving structure prediction accuracy from
unbiased MD simulations [29–32]. Herein, we used SAXS
information combined with Rosetta energy to identify those
models with the lowest possible scoring energy, and which
best fit the SAXS data [29]. Figure 2b shows the energies
of all models with respect to their SAXS discrepancy scores
(reduced χ2). The best-scored models display a similar
well-folded central core, faithfully matching the ab initio
envelope’s high-density region (Fig. 2c). The C-terminal
part containing the four conserved cysteine residues from
the 2Fe–2S cluster is less defined in the models, mostly
due to the high uncertainty in the iron-cluster geometry
and lack of structural constraints driving the modeling.
FhuF contains a 2Fe–2S cluster with unprecedented binding motif sequence Cys–Cys–x10–Cys–x2–Cys. Without a
template for the iron-cluster, predicting this region de novo
remains a challenging task in structural biology, even with
the incorporation of evolutionary coupling constraints [19].
Nevertheless, in most models, the cysteines are spatially
clustered, at a distance adequate to accommodate a 2Fe–2S
cluster (Fig. 3a). The well-defined core of the model is primarily α-helical, in agreement with the CD data, comprising
a four-helix bundle (α1:88–114; α2:153–177; α3:180–202;
α4:205–216) sandwiched between a three-stranded antiparallel β-sheet (β1:53–56; β2:125–129; β3:135–140) and a
two-stranded antiparallel β-sheet. The overall fold is analogous to the palm domain within AcsD/IcuA-like proteins
(Fig. 3b).
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319
Fig. 3 Relation of overall FhuF structural model with its closest
homologues. a Ribbon representation of the best-scored model. The
core four-helix bundle is colored in light blue, other helices in orange,
and beta-sheets in purple. The SG atoms of the cysteines known to
coordinate the 2Fe–2S centre at the C-terminal are depicted as yellow spheres. b Overlay of FhuF model and AcsD X-ray structure
(PDB:2W02) in transparent ribbon representation
Paramagnetic NMR suggests that the core
of the 2Fe2S cluster structure is similar
to that of other ferredoxins
properties of the 2Fe2S cluster by paramagnetic NMR spectroscopy, because it displays resolved features in both oxidation states. FhuF is unusual in this respect, because vertebrate 2Fe2S ferredoxins typically display unsuitable spectra
in the reduced state, whereas the opposite is observed for
2Fe2S plant ferredoxins [33]. Figure 4a shows the downfield region of 1D 1H NMR spectra of oxidized FhuF. The
spectrum of FhuF has a similar pattern of signals to that
observed in HuFd (Human ferredoxin), where a total of five
broad peaks (a–e) are observed, one at 11 ppm (e), and four
between 32 and 57 ppm (a–d) [33]. All five signals exhibit
anti-Curie temperature dependence (Fig. 4b), i.e., signals
shift further downfield as the temperature is increased, consistent with an antiferromagnetically coupled Fe(III)–Fe(III)
pair, as reported by EPR and Mössbauer spectroscopy
[11]. A 2Fe–2S cluster containing two antiferromagnetically coupled Fe(III) has an S = 0 ground-state which is
diamagnetic, thus the paramagnetic effect observed arises
from thermal population of excited spin-states in the orbital
manifold. The temperature dependence of these signals is
well reproduced by the VanVleck formalism using the Ei
The unprecedented binding motif of the 2Fe2S cluster,
involving cysteines 244, 245, 256 and 259 together with
the unusual gz value reported for the reduced form and the
unusual Mössbauer parameters for the ferric iron of the
reduced form of the cluster suggested the presence of a novel
2Fe–2S cluster structure [8, 11]. This is expected to affect
the electronic structure parameters of the cluster that can be
explored by NMR spectroscopy probing the paramagnetic
effect on the protons of the cysteines coordinating the cluster. Indeed, there are paramagnetically shifted signals both in
the oxidized and reduced forms of FhuF. In the reduced form
these shifts arise from the presence of an unpaired electron
in the 2Fe–2S cluster. In the oxidized form these arise from
the presence of thermally populated paramagnetic excited
states at room temperature even though the ground state of
the cluster is diamagnetic. Figures 4 and 5 show that FhuF
is amenable to detailed characterization of the electronic
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JBIC Journal of Biological Inorganic Chemistry (2021) 26:313–326
Fig. 4 Oxidised FhuF has a thermally populated paramagnetic
excited state. 1D 1H NMR spectra of oxidized FhuF (a). Temperature
dependence of hyperfine shift of β-CH2 and α-CH protons of oxidized
FhuF (b). Lines represent the fitting of Eq. (3) to the experimental
data, with Ai/h ranging from 3.07 MHz to 0.63 MHz using a single J
value of 300 cm−1 with a standard error of 2.5 cm−1
values obtained with J = 300 cm−1 and Aj/h ranging between
0.63 and 3.07 MHz. This J value is in the high range compared with those reported for spinach and algal ferredoxins
(J = 290 cm−1 and 185 cm−1) based on measurements of the
temperature dependence of paramagnetic shifts of cysteine
ligands to the clusters, the temperature dependence of magnetic susceptibility and on ENDOR, Mössbauer and EPR
data [34–36]. The Aj/h values for β-CH2 cysteine protons fall
in the expected range of 1 to 3 MHz obtained from proton
ENDOR data on a [ Fe4S4]3+ and rubredoxin models [21,
37]. The lower value of 0.63 MHz, calculated for peak e,
therefore, also argues for assigning this signal to an α-CH
proton of a cysteine bound to the cluster [21]. Therefore, by
analogy with HuFd, the peak at 11 ppm most likely accounts
for an α-hydrogen of one of the four ligated cysteines, and
the peaks between 57 and 32 ppm arise from the cysteinyl
β-hydrogens [33].
The 1D 1H NMR spectrum of reduced FhuF (Fig. 5a)
revealed nine peaks, five (A(III)–E(III)) between
199–104 ppm and four between 42 and 11 ppm (F(III)–I(II)).
Unlike the oxidized spectrum, the reduced spectrum of FhuF
does not resemble the spectra of reduced HuFd. Instead it
resembles the reduced spectrum of Anabaena 7120 vegetative
ferredoxin (Vfd) [38]. Despite the similar pattern, the signals
of the Anabaena ferredoxin do not reach beyond 140 ppm,
whereas in Fhuf, the most downfield signals reach to nearly
200 ppm. This suggests that one of the iron coordinating
cysteines has an unusually strong bond. Of the nine peaks,
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JBIC Journal of Biological Inorganic Chemistry (2021) 26:313–326
321
Fig. 5 Reduced FhuF localizes the electron in a specific iron. 1D
1
H NMR spectra of reduced FhuF (a). Temperature dependence
of hyperfine shift of β-CH2 and α-CH protons of reduced FhuF (b).
Lines represent the fitting of Eq. 3 to the experimental data, with Ai/h
ranging from 2.30 MHz to 0.31 MHz and using a single J value of
115 cm−1 with a standard error of 2.5 cm−1
the two less downfield-shifted signals exhibit anti-Curie
temperature dependence (H(II)-I(II)) while the remaining
seven, more far-shifted, exhibit Curie temperature dependence (A(III)-G(III)) (Fig. 5b). Signals with anti-Curie temperature dependence are assigned to protons of cysteines that
are ligated to Fe(II), whereas signals with Curie temperature dependence are assigned to protons of cysteines that are
ligated to Fe(III) [21, 33]. The observation of seven peaks
with Curie temperature dependence and the differences in
intensities of some peaks, for example peak F appears to be
of lower intensity, argue for the presence of heterogeneity
in the cluster environment in the reduced Fhuf. Indeed, the
observation of one clear extra peak from the 6 expected for a
pair of βCH2 and one αCH for each cysteine suggest that in
the reduced state there are at least two coordination modes
for the Fe (III). Nonetheless, Fig. 5b shows that the experimental data are well reproduced using J = 115 cm−1 and Aj/h
values between 0.31 and 2.13 MHz. Only peak C shows an
apparent temperature dependence that would require distinct
and unrealistically high values for Aj/h and J. Nonetheless,
the common J for all other signals together with a value of
Aj/h within the expected range of 1–3 MHz also places the
prediction of peak C in the correct chemical shift range. This
argues for the apparent steeper temperature dependence of
this signal to be of different origin, such as a temperature
dependent conformational change [39].
A J value of 115 cm−1 is also comparable with that
reported for spinach and algal ferredoxins (100 cm−1 and
13
322
115 cm−1) [35, 36]. The decrease in J value from oxidized to
reduced state has been proposed to be the result of the larger
ionic radius of Fe(II), which leads to a less efficient Heisenberg exchange mechanism between the two iron atoms [21].
Given that the J values for the oxidized and reduced states
of FhuF fall within typical values for 2Fe–2S ferredoxins,
and these values report on the coupling between the iron
atoms via the inorganic sulfurs, these results strongly suggest that the core of the cluster is maintained in FhuF [40].
Mössbauer data show that only the parameters for the Fe(III)
of the reduced cluster are unusual, whereas the parameters
for the Fe(II) are typical for tetrahedral coordination. This
is in agreement with paramagnetic 1H NMR experiments.
This observation and the fact that the differences in the cluster binding motif sequence of FhuF vs the typical binding
sequence for other ferredoxins is restricted to the separation
of the first two cysteines in the sequence (C–C–x10–C–x2–C
in FhuF vs the typical C–x2–C–x8-15–C–x2–C), we tentatively propose that the Fe(III) in the reduced state is bound
to cysteines 244 and 245. Binding of vicinal cysteines to iron
has precedent in the literature and was reported to lead to
Fig. 6 FhuF binds ferrichrome. 2D 1H15N TROSY-HSQC spectra
of 15N13C-labeled FhuF portraying the spectral changes observed
upon the addition of ferrichrome (a) and representative peak in fast-
13
JBIC Journal of Biological Inorganic Chemistry (2021) 26:313–326
a cysS-Fe-Scys angle that is systematically wider than the
ideal tetrahedral geometry, in agreement with the Mössbauer
data for FhuF [11, 41]. In the present case it appears to lead
also to different bonding strength by the two cysteines binding the Fe(III) in the cluster.
FhuF binds ferrichrome and apo‑ferrichrome
When oxidized, 2Fe–2S proteins are S’ = 0; however, some
paramagnetism arises from the population of the excited
states at room temperature. In FhuF, the effects of the
2Fe-2S paramagnetism are clearly reflected in the 1H15N
TROSY-HSQC of FhuF, where only 215 backbone peaks
out of 235 expected are observed (Figs. 1a, 6a). At least
20 peaks are undetected, and these most likely correspond
to the residues that fall in the “blind sphere”, the region
that surrounds the paramagnetic 2Fe–2S cluster [42]. Even
with the use of paramagnetic-tailored experiments it was
not possible to detect all the expected resonances [43, 44].
Nonetheless, upon the addition of ferrichrome to FhuF,
spectral changes were observed (Fig. 6a), including the
exchange regime with respective binding and fitting curve (b). R
refers to the concentration ratio between ligand and protein
JBIC Journal of Biological Inorganic Chemistry (2021) 26:313–326
disappearance of peaks. These results are consistent with
the binding of the Fe(III)-containing siderophore, which
being also paramagnetic, leads to fast relaxation and peak
broadening beyond detection in its vicinity [45]. Additionally, other spectral changes are observed, of which, peak
shifts in the fast-exchange regime allow the determination
of a dissociation constant of 53 ± 26 μM (Fig. 6b). Similarly,
upon the addition of apo-ferrichrome, spectral changes also
occur, suggesting the binding of apo-ferrichrome to FhuF.
Surprisingly, as previously found with ferrichrome but to a
lesser extent, some peaks are also bleached suggesting that
not only the presence of the paramagnetic center (Fe(III)
of ferrichrome) is contributing to this phenomenon (Figure
S2A). Given the expected presence of cavities in the structure (Fig. 2a), it is likely that the binding of both apo- and
holo-ferrichrome lead to conformational changes that bring
further regions of the protein into the reach of the “blind
sphere” generated by the 2Fe2S cluster.
323
Table 2 Reduction potentials
of FhuF as a function of
pH determined from cyclic
voltammetry measurements
pH
Formal
potential (V
vs SHE)
5.4
6.4
7.4
8.2
9.1
− 0.24 ± 0.07
− 0.30 ± 0.08
− 0.36 ± 0.09
− 0.42 ± 0.09
− 0.48 ± 0.09
FhuF does not display favorable electrochemical characteristics requiring extensive stabilization to observe a
redox signal from the adsorbed protein onto PGE (Fig. 7a
and Figure S3). This likely causes degradation of some of
the protein at the surface of the electrode giving rise to
the broad signal observed at higher potential in Fig. 7b,
which is not reproducible in terms of potential or linewidth
across different runs. By contrast, the signals at lower
potential are reproducible and display a half-height width
of the anodic and cathodic signals close to the theoretical
value of 90 mV for a single electron transfer step (Fig. 7b)
[46, 47]. The midpoint potential of – 370 mV vs SHE
at pH 7.4 is consistent with earlier studies which report
a reduction formal potential of − 310 ± 25 mV vs NHE
at pH 7.3 for FhuF at cryogenic temperatures [8]. Cyclic
voltammetry experiments performed at different pH values
(Table 2) show that FhuF presents a redox-Bohr effect as
also observed for a ferric-siderophore reductase from the
SIP subfamily from Shewanella frigidimarina [12].
The formal reduction potential changes by 60 mV for
every pH unit in good agreement with the expectation for
a Nernstian equilibrium for a coupled one proton and one
electron transfer (2.3RT/nF). Fitting of the pH dependence
of the potentials shows that the pkaox value must be lower
than 5 and the p ka red must be higher than 9 [48]. This
is especially significant, since the normal range of E.coli
growth can span pH 5 to 9 with some E.coli strains even
surviving in lower acidic environments [49]. Therefore,
the cyclic voltammetry data show that within this pH range
the FhuF formal reduction potential shifts from − 250 mV
to − 490 mV vs SHE. The lower value is particularly significant, since it broadens the diversity of siderophores
that can be reduced by FhuF and provides a rational for
the unexpected observation of Ferrioxamine B reduction
reported in the literature [8].
Fig. 7 FhuF displays pH dependent redox properties. Raw voltammograms of FhuF adsorbed onto PGE recorded at 100 mV/s in 20 mM
potassium phosphate buffer at pH 9.1 (blue), pH 8.2 (pink), pH 7.4
(black), pH 6.4 (green) and pH 5.4 (orange) (a) and Faradaic signal
of FhuF obtained by subtracting the capacitive current from the raw
voltammograms using QSoas software (b) [47]
The reduction potential of FhuF is pH dependent
broadening its catalytic capability
13
324
Conclusion
The FSR subclass of siderophore interacting proteins has
thus far resisted characterization. In this work we obtained
a structural and functional characterization of this important class of enzymes for iron uptake. FhuF binds both
ferrichrome and apo-ferrichrome, advocating for its role
as a bona-fide ferric siderophore reductase. The combination of SAXS data with the Rosetta derived model revealed
that FhuF shares the “palm domain” with the siderophore
biosynthetic enzymes, suggesting a common evolutionary origin that is distinct from that of SIP subfamily.
SIPs, despite performing the same function, display significant structural homology with the diverse family of
FAD/NAD(P)H oxidoreductases [50]. Paramagnetic NMR
spectroscopy established that the perturbation of the cluster vs typical 2Fe–2S ferredoxins appears to be confined
to the periphery involving only the cysteine ligands, in
particular those coordinating Fe(III) in the reduced state
showing the hallmarks of very different bond strengths
for the two cysteines coordinating this iron. This asymmetric bonding of one cysteine to Fe(III) may be at the
origin of the two conformations that appear to exist in
the reduced state. Nonetheless, the information that the
2Fe–2S rhomboid core is not disturbed combined with
the Mössbauer data and crystallographic knowledge of
the geometrical consequences of vicinal cysteine binding to iron allowed us to tentatively assign the ligands of
the two irons of the cluster. Knowing which of the two
irons is redox active in the cluster is essential to understand the molecular mechanism of siderophore reduction
by this protein once a more detailed structure is available. Interestingly, the presence of a redox-Bohr effect in
FhuF shows that this aspect of molecular mechanism is
common in siderophore-interacting proteins in both SIP
and the FSR subfamilies. For the proteins in the SIP subfamily, that use a flavin co-factor, this is not surprising
given the direct thermodynamic coupling of electron and
proton transfer in the isoalloxazine ring. By contrast for
proteins in the FSR subfamily it means that an acid–base
residue that titrates in the physiological pH range needs to
be placed near the 2Fe–2S cluster, eventually engaged in
H-bonding interaction, to allow proton-electron coupling
[51]. This strongly suggests that the pH dependence of the
reduction potential in siderophore interacting proteins is a
consequence of a common selective pressure on these proteins to enhance their physiological activity. Indeed, the
broader redox range afforded by the pH dependence of the
potential increases the diversity of siderophores that are
accessible for iron extraction by this protein and provides
a rational for the observation of reaction of FhuF with the
low potential siderophore ferrioxamine B [8]. The present
13
JBIC Journal of Biological Inorganic Chemistry (2021) 26:313–326
work sets the stage for a detailed investigation regarding
the mechanism of ferric-siderophore reduction together
with a detailed molecular characterization of the enzymes
responsible for this process.
Supplementary Information The online version contains supplementary material available at https://doi.org/10.1007/s00775-021-01854- y.
Acknowledgements The authors are grateful to Prof Alfred Trautwein,
who established the first contact with Prof Berthold Matzanke who
graciously made the FhuF expression system available that allowed
this work to be performed. ROL recalls vividly the ICBIC 5 organized in Lübeck by Alfred and which marked the start of his scientific
career. The authors are also grateful the Dr João Vicente for helpful
discussions and technical assistance with respect to the CD data. This
article is based upon work from COST Action CA15133, supported
by COST (European Cooperation in Science and Technology).We
acknowledge the France-Portugal PHC PESSOA program for support, project 40814ZE. Financial support was provided by European
EC Horizon2020 TIMB3 (Project 810856). This work was funded by
national funds through FCT– Fundação para a Ciência e a Tecnologia,
I.P. (FCT), Project MOSTMICRO-ITQB with refs UIDB/04612/2020
and UIDP/04612/2020. N-terminal sequencing service was provided
by the ITQB Research facilities. We acknowledge the use of BioSAXS beamline B21 (DLS-Dicot).The NMR data were acquired at
CERMAX, ITQB-NOVA, Oeiras, Portugal with equipment funded by
FCT, project AAC 01/SAICT/2016. IBT and GH are financially supported by national funds through the FCT PT-NMR PhD Program via
PD/BD/135187/2017 and PD/00065/2013, respectively. TNC is recipient of the grant CEECIND/01443/1017.
Compliance with ethical standards
Conflict of interest The authors declare no conflict of interests.
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long
as you give appropriate credit to the original author(s) and the source,
provide a link to the Creative Commons licence, and indicate if changes
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included in the article’s Creative Commons licence, unless indicated
otherwise in a credit line to the material. If material is not included in
the article’s Creative Commons licence and your intended use is not
permitted by statutory regulation or exceeds the permitted use, you will
need to obtain permission directly from the copyright holder. To view a
copy of this licence, visit http://creativecommons.org/licenses/by/4.0/.
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Authors and Affiliations
I. B. Trindade1 · G. Hernandez1 · E. Lebègue2 · F. Barrière3 · T. Cordeiro1 · M. Piccioli4,5 · R. O. Louro1
* R. O. Louro
louro@itqb.unl.pt
1
Instituto de Tecnologia Química e Biológica António Xavier
(ITQB‑NOVA), Universidade Nova de Lisboa, Av. da
República (EAN), 2780‑157 Oeiras, Portugal
2
Université de Nantes, CNRS, CEISAM UMR 6230,
44000 Nantes, France
3
Institut des Sciences Chimiques de Rennes‑UMR 6226,
Université Rennes, CNRS, 35000 Rennes, France
13
4
Department of Chemistry, Magnetic Resonance Center
(CERM), University of Florence, Via L. Sacconi 6,
50019 Sesto Fiorentino, Italy
5
Consorzio Interuniversitario Risonanze Magnetiche
di Metallo Proteine (CIRMMP), Via L. Sacconi 6,
50019 Sesto Fiorentino, Italy